ARTICLE IN PRESS

0038-0717/$ - se

doi:10.1016/j.so

�CorrespondE-mail addr

Soil Biology & Biochemistry 39 (2007) 2414–2419

www.elsevier.com/locate/soilbio

Short communication

A method for linking in situ activities of hydrolytic enzymes toassociated organisms in forest soils

Shufu Donga, Denise Brooksb,�, Melanie D. Jonesa, Susan J. Graystonb

aBiology and Physical Geography Unit, UBC Okanagan, 3333 University Way, Kelowna, BC, Canada V1V 1V7bDepartment of Forest Sciences, University of British Columbia, 2424 Main Mall, Vancouver, BC, Canada V6T 1Z4

Received 24 November 2006; received in revised form 5 March 2007; accepted 23 March 2007

Available online 4 May 2007

Abstract

A root window-based, enzyme-imprinted, membrane system has been modified to enable visualization of the activities of hydrolytic

enzymes (acid phosphatase, aminopeptidase, chitinase, and b-glucosidase) in situ in forest soils. The approach can be used to correlate

the distribution of enzyme activity with visible features such as roots, mycorrhizas, or mycelial mats. In addition, it enables accurate

spatial soil sampling for analysis of microbial communities associated with enzyme activities. The substrates are colorimetric conjugates

of napthol, where color develops instantly in the field, or fluorimetric conjugates of 4-methylumbelliferone, whose fluorescent products

are detected by a gel-documenting system. The method will allow important questions about the relationship between taxonomic and

functional diversity of soil microorganisms to be addressed and identification of enzyme activity hot-spots in soil.

r 2007 Elsevier Ltd. All rights reserved.

Keywords: Enzymes; Imprinting; Nutrient cycling; Roots; Root windows

1. Introduction

Enzymes have an obligatory role in catalyzing soilnutrient transformations (Burns and Dick, 2002). Mea-surement of soil enzyme activities has, therefore, beenrecommended as an extremely pertinent method formeasuring changes in soil quality (Dick, 1992), soilrecovery from disturbance or stress (Decker et al., 1999),and as the most appropriate indicator of microbialfunction (Caldwell, 2005). There are currently many well-utilized enzyme assays based on colorimetric and fluori-metric substrates that employ rapid microplate techniques,as reviewed by Caldwell (2005). However, these assays allinvolve soil sampling followed by lab analysis, inevitablyresulting in changes in enzyme activities (Tabatabai, 1994).Thus these methods, like those that probe for DNA andRNA of specific enzymes in soils (Kelly, 2003; Wellingtonet al., 2003), reveal only potential, not actual, enzymeactivity in soils.

e front matter r 2007 Elsevier Ltd. All rights reserved.

ilbio.2007.03.030

ing author. Tel.: +1778 888 3464; fax: +1 604 822 8645.

ess: denisebrooks3@hotmail.com (D. Brooks).

In their recent review of nitrogen cycling, Schimel andBennett (2004) argue that soil processes will only beunderstood if they are studied at much a finer scale than ispossible with conventional, destructive soil sampling. Here,we report novel in situ methods to detect hotspots of C, Nand P cycling activity in the soil profile. The methodsmodify and extend the field-based, root window approachof Grierson and Comerford (2000).

2. Method development

Root windows (transparent acrylic panel(77 cm� 52 cm� 0.6 cm) with a 30 cm� 30 cm trap door(Grierson and Comerford 2000)) were installed in a rangeof Douglas-fir (Pseudotsuga menziesii) stands in the fieldfor 5 months prior to imprinting. A membrane of eitherchromatography (Whatman, Cat No. 3030-861) or filterpaper (Whatman, Cat. No. 1001 055), treated with either amixture of substrate and colorimetric reagent or afluorimetric substrate, was placed directly on the soilsurface and enzyme activity detected by the appearance ofeither colored or fluorescent products on the membrane.Optimal duration of imprinting was 30min for all enzymes

ARTICLE IN PRESS

Fig. 1. (a) Color or fluorescence development, expressed as gray value, of imprints from assays for acid phosphatase, aminopeptidase, chitinase, and b-glucosidase, with increasing time of contact with rhizobox soil (data from two runs for each enzyme). Note that increasing gray value represents higher

levels for the fluorescent products of chitinase and b-glucosidase; whereas higher levels of the colored products of phosphatase and aminopeptidase result

in lower gray values. (b) Fluorescence of 4-methylumbelliferone residue, expressed as gray scale, adhering to untreated pieces of filter paper applied to

rootboxes containing Douglas-fir seedlings 1–8 days after initial assays (day 0) for chitinase and b-glucosidase activity. As fluorescence increases, gray

value increases. Mean values7SEM of three replicate rhizoboxes containing 6-months old. Douglas-fir seedlings growing in field soil.

S. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–2419 2415

except phosphatase, which required 60min (Fig. 1a). Afterexposure, the imprints were carefully removed, rinsed withdeionized water, air-dried and scanned. Membranes werehandled throughout with latex gloves or sterilized forceps.

To detect acid phosphatase activity, chromatographypaper was soaked for 1min in a 1:10 (v/v) mixture offreshly prepared 50mM a-naphthyl phosphate (SigmaN7255) and 10mM Fast Red TR (Sigma F2768), bothprepared in 50mM pH 5.6 citrate buffer (Dinkelaker andMarschner 1992), and then air-dried. Standards (SigmaP3627, from wheat germ) of 0–0.35 enzyme units

(EU)ml�1 in 5 ml pH 5.6 citrate buffer were applied toseparate pieces of membrane and placed adjacent to testmembranes on soil surfaces The intensity of purple-redcolor after conversion to gray scale in Adobe PhotoshopElements 2.0, represented acid phosphatase activity(Fig. 2a). Control membranes treated with only Fast Redshowed no color after imprinting.Membranes to detect aminopeptidase activity were

prepared by soaking in 20mM L-leucyl 2-naphthylamide(Sigma L0376, prepared in 95% alcohol) followed by air-drying. Fast Blue BB (2.4mM in DI water, Sigma F0250)

ARTICLE IN PRESS

Fig. 2. Imprints and soil profiles from root windows in interior Douglas-fir stands near Barriere, British Columbia, Canada. (a) phosphatase imprint; (b)

aminopeptidase imprint, (c) soil image overlain with the same imprint, (d) image of soil profile; (f) b-glucosidase imprint and (e)associated soil image; (h)

chitinase imprint and (g) associated soil image. All images are at the same scale.

S. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–24192416

was applied in a fine mist after imprints and standards wereremoved from soil surfaces. Imprints were then exposed to150W infrared light (1min) to minimize development ofnon-specific background color (Humble et al., 1977). Anorange-red color represented aminopeptidase activity.Standards for the aminopeptidase assay were preparedfrom a fungal protease/peptidase complex of Aspergillus

oryzae (Sigma, P6110) and applied to membranes in 5 mlaliquots containing 0, 3.9, 7.8, 15.6, and 31.2 EUml�1.Control membranes received no substrate and exhibited nocolor when sprayed with Fast Blue. Although the resolu-tion of aminopeptidase activity was lower than acidphosphatase, the association of aminopeptidase activitywith roots could be clearly observed (Fig. 2b–d).

Chitinase activity was visualized on membranes soakedin 5mM 4-methylumbelliferyl-N-acetyl-b-glucosaminide(Sigma M2133) in 2-methoxylethanol (Sigma M5378).Membranes for b-glucosidase used 4-methylumbelliferyl-b-glucopyranoside dehydrate (Sigma M3633) as a sub-strate. Activity of these enzymes on these substrates releasefluorescent 4-MUB (Hoppe, 1983; Pritsch et al., 2004). Inthe lab imprints were imaged with a gel documentationsystem (Gel LOGIC 440, Eastman Kodak Company, NewHaven, CT, USA). Persistence of fluorescent residues onsoil surfaces was found for 8 days following imprinting(Fig. 1b), so we recommend that tests using 4-MUB-linked

substrates be at least 10 days apart. 4-MUB (SigmaM1381) standards of 0, 0.078, 0.156, 0.313, and0.625mM in 2-methoxylethanol were applied in 5 mlaliquots to membranes to generate standard curves forboth chitinase and b-glucosidase activity. When testmembranes prepared with either the substrate for chitinaseor the substrate for b-glucosidase were applied to soilsurfaces, fluorescence could be detected on the resultingimprints (Fig. 2e–h). The boundaries of the fluorescentspots were diffuse but activities could be associated withspecific structures in the soil. Control membranes exhibitedlittle fluorescence, with gray values typically below 100.

3. Applications of the method

By generating standard curves either from enzymestandards applied to test membranes in the field (phos-phatase and aminopeptidase) or from known amount ofproduct applied to membranes just before imaging (MUfor chitinase and b-glucosidase), it is possible to use themethod to semi-quantitatively estimate in situ enzymeactivities. For example, imprints taken at root windowsinstalled in a mixed Picea engelmannii/Abies lasiocarpa

stand (Hagerman et al., 1999) detected significantly higherphosphatase activities in nine natural stands (24.472.1)than in nine clear cuts (6.572.2). The method also allows

ARTICLE IN PRESS

Fig. 3. Phosphatase activity derived from scanned 20 cm� 20 cm imprints taken from a Douglas-fir/birch chronosequence of four ages: young (3–6 years),

canopy closure (24–27 years), stem exclusion (55–60 years), and older (88–100 years) near Enderby, British Columbia, Canada (mean values7SEM for

total number of active areas; n ¼ 3). Different letters indicate a significant difference at Po0.05 according to Tukey test for the total number of active

areas illustrated in the bar. (a) Total number of visible phosphatase-active areas per imprint. (b) Frequency distribution of different sizes of phosphatase-

active areas larger than 50 pixels in size on scanned imprints.

S. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–2419 2417

determination of the extent and spatial distribution ofenzyme activity. For example, the observation that thenumber, average size, and intensity of phosphatase hotspots increased with stand age in mixed P. menziesii/Betula

papyrifera stands (Fig. 3a and b), prompted us todetermine whether the composition of the microbialcommunity associated with high phosphatase patcheschanges with stand age.

A major strength of the in situ methods describedhere is that they can be used to correlate enzyme activitieswith plant, fungal, and bacterial communities in soils priorto destructive sampling. The location of macroscopic

features, such as soil horizons, roots, and mycelial fans,can be recorded with high-resolution digital photography.These images can then be overlain with scans of theimprints, and the two images aligned using holes in theimprints created by pins inserted through the root window(Grierson and Comerford, 2000). Such an approachhas demonstrated that, in clear cut soils, b-glucosidaseand chitinase activities were found almost exclusivelyin association with roots and decaying material,whereas in adjacent Douglas-fir stands, activity wasmainly associated with fungal mats or organic horizons(Fig. 4a and b).

ARTICLE IN PRESS

Fig. 4. Percentage of chitinase and b-glucosidase activity associated with organic soil (OS), mineral soil (MS), roots (RT), fungal mats (FM), and decayed

material (DM) on imprints taken in three root windows per site in (a) Douglas-fir forests and (b) clear cuts near Barriere, British Columbia, Canada (mean

values7SEM; n ¼ 4 sites).

S. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–24192418

By using molecular techniques such as DGGE orT-RFLP, soil fungal and prokaryotic communities asso-ciated with hot spots of enzyme activity can be described.To do this, the imprints can be used to create templatesfrom clear acetate sheets for sampling soils at mm scales.This approach will complement approaches that probe forspecific genes (Kelly, 2003; Wellington et al., 2003) bycorrelating the presence of organisms with directlymeasured activity.

Many factors that affect enzyme activity, such as pH,moisture and temperature, will vary at a fine scale across

individual root windows. Although this variation might beseen as a disadvantage by researchers used to measuringbulk soil enzyme activities, we see it as a major advantageof the method. This method gives us confidence that we aredetecting actual enzyme activity as it occurs in the field insoil microsites. This fine-scale variation in environmentalconditions is lost during typical soil sampling. Thus,this is exactly the type of approach that will allow us tostudy soils at the scales suggested by Schimel and Bennett(2004) and hence, deepen our understanding of soilprocesses.

ARTICLE IN PRESSS. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–2419 2419

Acknowledgements

The authors gratefully acknowledge financial supportfrom the Special Research Opportunity and DiscoveryGrant programs of the Natural Sciences and EngineeringResearch Council of Canada, the Forest Science Programof the Forest Investment Account of British Columbia, andthe award of a University of British Columbia Cordula andGunter Paetzold Fellowship to DB. We are grateful forexcellent field assistance from Jason Barker, Anne Bern-hardt, Ben Chester, Julie Deslippe, Alanna Leverrier,Kristen Mackay, Adam McCaffrey, Chelsea Ricketts,and Jeff Sherstobitoff. This research was based on initialdiscussions and considerable advice from Pauline Griersonduring a sabbatical leave by MJ at the University ofWestern Australia.

References

Burns, R.G., Dick, R.P., 2002. Enzymes in the Environment: Activity,

Ecology and Applications. Marcel Dekker, New York, 614pp.

Caldwell, B.A., 2005. Enzyme activities as a component of soil

biodiversity: a review. Pedobiologia 49, 637–644.

Decker, K.L.M., Boerner, R.E.J., Morris, S.J., 1999. Scale-dependent

patterns of soil enzyme activity in a forested landscape. Canadian

Journal of Forest Research—Revue Canadienne De Recherche

Forestiere 29, 232–241.

Dick, R.P., 1992. A review—long-term effects of agricultural systems on

soil biochemical and microbial parameters. Agriculture Ecosystems

and Environment 40, 25–36.

Dinkelaker, B., Marschner, H., 1992. In vivo demonstration of acid-

phosphatase-activity in the rhizosphere of soil-grown plants. Plant and

Soil 144, 199–205.

Grierson, P.F., Comerford, N.B., 2000. Non-destructive measurement of

acid phosphatase activity in the rhizosphere using nitrocellulose

membranes and image analysis. Plant and Soil 218, 49–57.

Hagerman, S.M., Jones, M.D., Bradfield, G.E., Gillespie, M., Durall,

D.M., 1999. Effects of clear-cut logging on the diversity and

persistence of ectomycorrhizae at a subalpine forest. Canadian Journal

of Forest Research—Revue Canadienne De Recherche Forestiere 29,

124–134.

Hoppe, H.G., 1983. Significance of Exoenzymatic Activities in the

Ecology of Brackish Water - Measurements by Means of Methylum-

belliferyl-Substrates. Marine Ecology-Progress Series 11, 299–308.

Humble, M.W., King, A., Phillips, I., 1977. Api zym—simple rapid system

for detection of bacterial enzymes. Journal of Clinical Pathology 30,

275–277.

Kelly, J.J., 2003. Molecular techniques for the analysis of soil microbial

processes: functional gene analysis and the utility of DNA micro-

arrays. Soil Science 168, 597–605.

Pritsch, K., Raidl, S., Marksteiner, E., Blaschke, H., Agerer, R., Schloter,

M., Hartmann, A., 2004. A rapid and highly sensitive method for

measuring enzyme activities in single mycorrhizal tips using

4-methylumbelliferone-labelled fluorogenic substrates in a microplate

system. Journal of Microbiological Methods 58, 233–241.

Schimel, J.P., Bennett, J., 2004. Nitrogen mineralization: challenges of a

changing paradigm. Ecology 85, 591–602.

Tabatabai, M.A., 1994. Soil enzymes. In: Anonymous Microbiological

and Biochemical Properties. Soil Science Society of America, Inc., pp.

775–833.

Wellington, E.M.H., Berry, A., Krsek, M., 2003. Resolving functional

diversity in relation to microbial community structure in soil:

exploiting genomics and stable isotope probing. Current Opinion in

Microbiology 6, 295–301.